long-term continuous electroencephalography monitoring in a rodent pup

Long-Term Continuous Electroencephalography Monitoring in a Rodent Pup

Transfer a single implanted animal to a recording chamber. Only one implanted animal can be monitored at a time, but other non-implanted animals can be with it.
To set up the recorder, connect the power supply to the receiver base. Next, connect the base to a data acquisition system via BNCs. Now, place the cage on the signal receiver base. The signal light should illuminate. Data can now be recorded.
First, connect the receiver base to an A/D converter, and connect the converter to a computer. Using the software settings, set the sample rate of the recording to 500 per second. The transmitter sends EEG signals between 0.1 to 100 Hertz with an 8 decibels per octave roll-off at either end.
Setting an appropriate sample rate that is above Nyquist is critical. We recommend 500 samples per second.
While the animals are being recorded from, watch for signs of stress and dehydration. Pups should not be in the chamber for more than two hours.

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All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Surgical Preparation
<ol>
	<li>Clean and prepare the transmitter to ensure safe and sterile surgery. Remove the transmitter from its anti-static packaging and either spray or soak in 70% ethanol. Rinse transmitter with sterile saline and place between sterile cotton sponges soaked in sterile saline or keep submerged in sterile saline.</li>
	<li>Collect and sterilize the tools required for surgery, steam autoclave for sterilization. See table of materials and reagents for the list of surgical tools.</li>
</ol>
2. Surgical Implantation
<ol>
	<li>Anesthetize animals and maintain anesthesia according to IACUC-approved protocol. At initiation and during surgery check the toe-pinch reflex every 15 min. The lack of response indicates sufficient level of anesthesia.
	<ol>
		<li>For pups, use anesthesia by isoflurane (4%) with O2 (100%). For adults, use ketamine (100 mg/kg) with xylazine (10 mg/kg).</li>
	</ol>
	</li>
	<li>Fix position in stereotaxic frame. Place the ear bar tips in the auditory meatus. Do not excessively tighten ear bars as the skull is very soft in young rat pups. Secure the anesthesia nose cone.
	<ol>
		<li>Keep the animal warm during surgery by placing it on the heating pad set to 37 &#176;C. In adult animals, apply lubricating ointment to the eyes of the animal.</li>
	</ol>
	</li>
	<li>Sterilize incision site and maintain sterile surgical field.
	<ol>
		<li>Swab the scalp with alternating applications of 70% ethanol and betadine. Start in the center of the scalp and make increasingly wider concentric circles.</li>
		<li>Cover the animal with drape and conduct the surgery over draped animal. Maintain the sterile surgical field by lining the surgical set-up with sterile drapes, spray equipment with 70% ethanol.</li>
		<li>Wear sterile surgical gloves and a gown (or as required by the institution). To help maintain a sterile field, use a surgical assistant.</li>
	</ol>
	</li>
	<li>Make an incision on the scalp of the animal slightly behind the eyes along the midline, approximately 2 cm. Use caution when inserting the scalpel as the skull is still very soft in young rat pups. Make a single cut so the incision bleeds less and heals faster.</li>
	<li>Expose the skull. Prepare a clean and dry area to maximize the bond between the transmitter and the bones of the skull. Use aneurism clips to grasp scalp.
	<ol>
		<li>Gently pull scalp away from midline at four corners. Look for anatomical landmarks such as bregma and lambda in the skull. Remember skull bones are not fused in animals at this age. Use the Paxinos atlas of stereotaxic coordinates to find the correct location for the burr hole.</li>
	</ol>
	</li>
	<li>Use a Dremel-type tool with a burr-type drill bit. Create two burr holes in desired recording positions with the holes being bigger than 300 &#181;m in diameter. Place the burr hole for the reference electrode over the cerebellum behind the lambda of the skull.</li>
	<li>Ensure that the wires on the transmitter are aligned with the burr holes. If the electrode wires are not aligned, glue contamination of the electrodes is likely, and will result in poor signal. To align the wires, check the fit of the transmitter and gently bend electrodes to line up over the intended sites for burr holes.</li>
	<li>Trim electrode leads. Use surgical scissors to trim the electrodes to the desired length. The electrode depth is important for the type of recording required for the experiment (i.e., place the electrodes above dura for EEG recordings, or use stereotaxic coordinates for defined brain structures).</li>
	<li>Liberally apply cyanoacrylate on the base of the transmitter to cover the area making sure to avoid coating the electrodes. Cyanoacrylate glue is an electrical insulator, contaminating electrodes with glue will result in no signal.
	<ol>
		<li>If recording from deep brain structures, mount the transmitter on the cannula holder and place it in the stereotaxic arm for z-axis control. Lower the transmitter using the stereotaxic arm to appropriate depth and place cyanoacrylate gel around the transmitter.</li>
	</ol>
	</li>
	<li>Thoroughly dry skull before placing transmitter to ensure strong adhesive bond. Apply transmitter coated with cyanoacrylate to the skull. Take care to align electrodes with corresponding burr holes.
	<ol>
		<li>Try to avoid damaging major vascular structures. Hold the transmitter in place with slight pressure for one minute. Use slight pressure to form a strong bond between the transmitter and the skull.</li>
	</ol>
	</li>
	<li>Apply additional cyanoacrylate, enough to completely seal transmitter/skull interface. To ensure a good fit and strong bond, maximize the surface area of the glue that contacts the skull. Apply the cyanoacrylate adhesive in a circle around the transmitter, making sure both skull and the wall of the transmitter are covered.</li>
	<li>Apply chemical accelerant (0.1 ml) through a syringe around the cyanoacrylate at the base of the implanted transmitter. Use accelerant sparingly, taking care not to apply to adjacent tissue. 
	Note: Chemical acceleration of the cyanoacrylate curing ensures that the strong bond between the transmitter and the skull is formed quickly. Cyanoacrylate accelerant is useful to speed curing of adhesive but is not necessary.</li>
	<li>Remove the accelerant by washing the area thoroughly with sterile saline. Cyanoacrylate accelerator may cause tissue irritation if not washed from the area of the incision. To wash the area, fill a 1.0 ml syringe with sterile saline and irrigate the area through a syringe needle. Generally, 0.5 ml of saline is enough to wash out the accelerator.</li>
	<li>Suture the skin around the base of the transmitter, but do not cover the transmitter. The top of transmitter must be above skin to efficiently transmit neural signals. Skin should be reasonably tight around the transmitter and the glue around the unit. Use Vicryl or silk suture (soft thread); skin in immature animals is soft and is easily damaged if soft sutures are not used. For adult animals, use any suturing material.</li>
	<li>Remove animal from stereotaxic frame and place on heated blanket for recovery.</li>
	<li>Ensure animals are warm (37 &#176;C) and ambulatory (i.e., completely recovered) before returning to the dam. Ensure that the animal is hydrated by pinching the skin on the animal&#39;s back (if the animal is dehydrated, the skin will remain deformed). If animal is dehydrated, administer sub-cutaneous injection of lactated Ringer&#39;s buffer. Do not leave the animal unattended until it has regained sufficient consciousness to maintain sternal recumbency.
	<ol>
		<li>Administer buprenorphine (0.05 mg/kg) to animals for post-surgical pain management and a sub-cutaneous injection of 0.1 ml bupivacaine around the injection site. 
		Note: From start to finish the entire procedure should be completed in 5-10 min for animals of this age (postnatal day 6). Surgical time may take longer for older animals.</li>
	</ol>
	</li>
</ol>
3. Care and Housing
Note: Some dams may not tolerate pups implanted with the device. Dams may need to be selected who are tolerant. It is acceptable for the dam to move pups around the cage by picking them up by the transmitter.
<ol>
	<li>Once animals are weaned, singly-house them to avoid the removal of the devices from their cage mate.</li>
	<li>Euthanize animals by lethal dose of pentobarbital (25 mg/kg) or isoflurane (in a bell jar) when signs of distress are present.</li>
	<li>Note: some animal housing cages with wire inserts may interfere with the implanted transmitters.&#160; Be sure to check the height of the wire insert to make sure that animals cannot get the transmitter caught between the &#39;bars&#39; of the wire insert.</li>
</ol>
4. Recording EEG
<ol>
	<li>Place the animal in a cage by itself or co-housed with littermates and the dam. However, place only one implanted animal in a single cage. Do not leave pups alone in the recording chamber for more than 2 hr. Monitor the animals for signs of distress and dehydration.</li>
	<li>Connect the provided power supply to the receiver base and verify the power light is illuminated. Connect the receiver base to a data acquisition system using Bayonet Neil-Concelman (BNC) cables.</li>
	<li>Place the animal cage on top of the receiver base (Figure 1). The &#34;signal&#34; light should illuminate indicating a transmitter has been detected. Data may now be recorded.</li>
	<li>To record data, connect the receiver base to an analogue-to-digital converter and connect the converter to a computer (Figure 2).</li>
	<li>Set the sampling rate of the recording. Ensure the data is sampled properly. Select at least 250 Hz sampling rate (500 Hz recommended) for recording (bandwidth of the transmitter is 0.1-100 Hz).</li>
	<li>Save digitized data and analyze using signal-processing software packages such as Matlab.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sterile Surgical Gloves</td>
			<td>Protective Industrial Products</td>
			<td>100-3201 PF</td>
			<td>Powder Free Sterile Latex Surgical Glove</td>
		</tr>
		<tr>
			<td>Scalpel Handle</td>
			<td>FST</td>
			<td>10003-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel Blade #15</td>
			<td>FST</td>
			<td>10015-00</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Scissors</td>
			<td>FST</td>
			<td>14090-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Burr tool</td>
			<td>Ram Products, Inc.</td>
			<td>Microtorque II</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine burr</td>
			<td>FST</td>
			<td>19007-07</td>
			<td></td>
		</tr>
		<tr>
			<td>Aneurism clip</td>
			<td>ROBOZ</td>
			<td>RS-5422</td>
			<td></td>
		</tr>
		<tr>
			<td>Toothed Forceps</td>
			<td>FST</td>
			<td>11022-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton-Tipped applicators</td>
			<td>McKesson</td>
			<td>24-103</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle Driver</td>
			<td>WPI</td>
			<td>521725</td>
			<td>Olsen-Hegar Needle Holder</td>
		</tr>
		<tr>
			<td>Cyanoacrylate gel</td>
			<td>Henkel</td>
			<td>Loctite 4541</td>
			<td></td>
		</tr>
		<tr>
			<td>Cyanoacrylate accelerant</td>
			<td>Henkel</td>
			<td>Loctite 7452</td>
			<td></td>
		</tr>
		<tr>
			<td>Suture</td>
			<td>Ethicon</td>
			<td>Vicryl RB-1 J304</td>
			<td></td>
		</tr>
		<tr>
			<td>Elecrocautery disposable</td>
			<td>Bovie</td>
			<td>AA01</td>
			<td>Fine Tip</td>
		</tr>
		<tr>
			<td>Surgical Tray</td>
			<td>FST</td>
			<td>20311-21</td>
			<td></td>
		</tr>
		<tr>
			<td>Epitel Receiver Base</td>
			<td>Epitel Inc</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Epitel wireless transmitter</td>
			<td>Epitel Inc</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Biopac digitizer</td>
			<td>Biopac</td>
			<td>MP-150</td>
			<td></td>
		</tr>
		<tr>
			<td>PC-compatible computer</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

Source: Zayachkivsky, A. et al. <a href="https://app.jove.com/v/52554">Long-term Continuous EEG Monitoring in Small Rodent Models of Human Disease Using the Epoch Wireless Transmitter System.</a> J. Vis. Exp. (2015)
The video demonstrates long-term continuous electroencephalography (EEG) monitoring in rodent pups using a wireless transmitter system. The transmitter contains two recording electrodes and one reference electrode extending to the brain&#39;s surface. Upon placing the pup in a recording chamber, EEG can be continuously monitored throughout the pup&#39;s development into adulthood.

<img alt="Figure 1" src="/files/ftp_upload/22864/22864_Figure1.jpg" /> 
Figure 1: The transmitter and receiver. This particular wireless transmitter (A) weighs 4 g and displaces &#60;1.4 cm3 of volume and with a footprint of 7 x 12 mm is easily mounted to the skull of rats and mice. The transmitter can amplify 2 channels of biopotentials for up to 6 months after which the battery is drained. Larger batteries can be used for longer recording time. Ani...

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

Take a rodent pup with a wireless electroencephalogram or EEG transmitter implanted in its skull.
The transmitter contains electrodes that extend to the brain&#39;s surface. Two recording electrodes monitor the electrical activity of neurons, and one reference electrode helps reduce signal noise.
Place a transmitter-implanted pup in a cage inside a recording chamber. The chamber contains a wireless receiver base connected to a data acquisition system that receives signals wirelessly.
Place non-implanted pups in the cage to reduce stress from isolation for the transmitter-implanted pup and start acquiring signals.
Neurotransmitters released at the synapses bind to receptors on postsynaptic neurons, inducing an ion influx that changes the membrane potential, termed postsynaptic potential.
The electrodes record postsynaptic potentials from a large number of neurons extracellularly. The summed synaptic activity of neighboring neurons is recorded as EEG signals.
The miniature transmitter allows continuous monitoring of EEG signals throughout the pup&#39;s development into adulthood.

13 Views. Long-Term Continuous Electroencephalography Monitoring in a Rodent Pup

Video: Long-Term Continuous Electroencephalography Monitoring in a Rodent Pup - Experiment

Simultaneous Video-EEG-ECG Monitoring to Identify Neurocardiac Dysfunction in Mouse Models of Epilepsy

In epilepsy, seizures can evoke cardiac rhythm disturbances such as heart rate changes, conduction blocks, asystoles, and arrhythmias, which can potentially increase risk of sudden unexpected death in epilepsy (SUDEP). Electroencephalography (EEG) and electrocardiography (ECG) are widely used clinical diagnostic tools to monitor for abnormal brain and cardiac rhythms in patients. Here, a technique to simultaneously record video, EEG, and ECG in mice to measure behavior, brain, and cardiac activities, respectively, is described. The technique described herein utilizes a tethered (i.e., wired) recording configuration in which the implanted electrode on the head of the mouse is hard-wired to the recording equipment. Compared to wireless telemetry recording systems, the tethered arrangement possesses several technical advantages such as a greater possible number of channels for recording EEG or other biopotentials; lower electrode costs; and greater frequency bandwidth (i.e., sampling rate) of recordings. The basics of this technique can also be easily modified to accommodate recording other biosignals, such as electromyography (EMG) or plethysmography for assessment of muscle and respiratory activity, respectively. In addition to describing how to perform the EEG-ECG recordings, we also detail methods to quantify the resulting data for seizures, EEG spectral power, cardiac function, and heart rate variability, which we demonstrate in an example experiment using a mouse with epilepsy due to Kcna1 gene deletion. Video-EEG-ECG monitoring in mouse models of epilepsy or other neurological disease provides a powerful tool to identify dysfunction at the level of the brain, heart, or brain-heart interactions.

Here, we present a protocol to record brain and heart bio signals in mice using simultaneous video, electroencephalography (EEG), and electrocardiography (ECG). We also describe methods to analyze the resulting EEG-ECG recordings for seizures, EEG spectral power, cardiac function, and heart rate variability.

In epilepsy, seizures can evoke cardiac rhythm disturbances such as heart rate changes, conduction blocks, asystoles, and arrhythmias, which can ...

JoVE Journal

Genetics

Use of a Wireless Video-EEG System to Monitor Epileptiform Discharges Following Lateral Fluid-Percussion Induced Traumatic Brain Injury

The lateral fluid percussion injury (FPI) model is well established and has been used to study TBI and post-traumatic epilepsy (PTE). However, considerable variability has been reported for the specific parameters used in different studies that have employed this model, making it difficult to harmonize and interpret the results between laboratories. For example, variability has been reported regarding the size and location of the craniectomy, how the Luer lock hub is placed relative to the craniectomy, the atmospheric pressure applied to the dura and the duration of the pressure pulse. Each of these parameters can impact injury severity, which directly correlates with the incidence of PTE. This has been manifested as a wide range of mortality rates, righting reflex times and incidence of convulsive seizures reported. Here we provide a detailed protocol for the method we have used to help facilitate harmonization between studies. We used FPI in combination with a wireless EEG telemetry system to continuously monitor for electrographic changes and detect seizure activity.&#160; FPI is induced by creating a 5 mm craniectomy over the left hemisphere, between the Bregma and Lambda and adjacent to the lateral ridge. A Luer lock hub is secured onto the skull over the craniectomy. This hub is connected to the FPI device, and a 20-millisecond pressure pulse is delivered directly to the intact dura through pressure tubing connected to the hub via a twist lock connector. Following recovery, rats are re-anesthetized to remove the hub. Five 0.5 mm, stainless steel EEG electrode screws are placed in contact with the dura through the skull and serve as four recording electrodes and one reference electrode. The electrode wires are collected into a pedestal connector which is secured into place with bone cement. Continuous video/EEG recordings are collected for up to 4 weeks post TBI.

Here we present a protocol to induce severe TBI with the lateral fluid percussion injury (FPI) model in adult, male Wistar rats. We also demonstrate the use of a wireless telemetry system to collect continuous video-EEG recordings and monitor for epileptiform discharges consistent with post-traumatic epileptogenesis.&#160;

The lateral fluid percussion injury (FPI) model is well established and has been used to study TBI and post-traumatic epilepsy (PTE). However, ...

Behavior

Simultaneous Monitoring of Wireless Electrophysiology and Memory Behavioral Test as a Tool to Study Hippocampal Neurogenesis

Brainwaves amplitude obtained from electroencephalography (EEG) has been well-recognized as a basis for cognitive capacity, memory, and learning on animals and humans. Adult neurogenesis mechanism is also linked to memory and learning improvement. Traditionally, researchers used to assess learning and memory parameters in rodent models by behavioral tasks. Therefore, the simultaneous monitoring of behavioral changes and EEG is particularly interesting in correlating data between brain activity and task-related behaviors. However, most of the equipment required to perform both studies are either complex, expensive, or uses a wired setup network that hinders the natural animals&#8217; movement. In this study, EEG was recorded with a wireless electrophysiology device along with the execution of a novel object recognition task (NORT). The animal&#8217;s behavior was monitored simultaneously by a video tracking system. Both recordings were analyzed offline by their timestamps which were synched to link EEG signals with the animal&#8217;s actions. Subjects consist of adult Wistar rats after medium-term environmental enrichment treatment. Six skull screw electrodes were fixed in pairs on both hemispheres over frontal, central, and parietal regions and were referenced to an electrode located posterior of the nasal bone. NORT protocol consists of exposing the animal to two identical objects for 10 min. After 2 h and 24 h, one of the objects was replaced with a novel one. Exploration time for each object was monitored by a behavioral tracking software (BTS) and EEG data recording. The analysis of the EEG synced with behavioral data consists of estimations of alpha and beta relative band power and comparisons between novel object recognition versus familiar object exploration, between three experimental stages. In this manuscript, we have discussed electrodes manufacturing process, epidural electrodes implantation surgery, environmental enrichment protocol, NORT protocol, BTS setup, EEG &#8211; BTS coupling for simultaneous monitoring in real-time, and EEG data analysis based on automatic events detection.

The protocol presented here provides information on the simultaneous electroencephalography (EEG) and behavioral assessment in real-time. We have discussed all steps involved in this protocol as an attractive solution for researchers in many fields of neuroscience, particularly in learning and memory areas.

Brainwaves amplitude obtained from electroencephalography (EEG) has been well-recognized as a basis for cognitive capacity, memory, and learning on ...

Long-Term Continuous Electroencephalography Monitoring in a Rodent Pup

Transcript